Simultaneous stabilization of actin cytoskeleton in multiple nephron-specific cells protects the kidney from diverse injury

Chronic kidney diseases and acute kidney injury are mechanistically distinct kidney diseases. While chronic kidney diseases are associated with podocyte injury, acute kidney injury affects renal tubular epithelial cells. Despite these differences, a cardinal feature of both acute and chronic kidney diseases is dysregulated actin cytoskeleton. We have shown that pharmacological activation of GTPase dynamin ameliorates podocyte injury in murine models of chronic kidney diseases by promoting actin polymerization. Here we establish dynamin’s role in modulating stiffness and polarity of renal tubular epithelial cells by crosslinking actin filaments into branched networks. Activation of dynamin’s crosslinking capability by a small molecule agonist stabilizes the actomyosin cortex of the apical membrane against injury, which in turn preserves renal function in various murine models of acute kidney injury. Notably, a dynamin agonist simultaneously attenuates podocyte and tubular injury in the genetic murine model of Alport syndrome. Our study provides evidence for the feasibility and highlights the benefits of novel holistic nephron-protective therapies.

T he leading causes of acute kidney injury (AKI) are ischemia, hypoxia, or nephrotoxicity 1 . While it can be reversed, AKI represents a significant healthcare problem with high mortality and no definitive treatment. Regardless of its etiology, AKI primarily injures polarized epithelial cells of the renal tubules whose apical microvilli form the tubular brush border that participates in coordinating essential electrolyte and water transport 2 . An early morphological feature of AKI is loss of the brush border and cell polarity due to the breakdown of the actomyosin cortex at the apical membrane 1 .
The establishment and maintenance of cell polarity involve signaling cascades, membrane trafficking, and cytoskeletal dynamics, all of which are highly coordinated 3 . The organization of the apical membrane is largely determined by the architecture of the actomyosin networks 4 , which establishes cortex stiffness, thus facilitating the clustering of polarity proteins. While myosin II motors are considered the primary generator of cortical stiffness 5,6 , the architecture of the actomyosin cortex is established by a myriad of actin-binding proteins (ABPs) 7 .
In addition to known ABPs, the brush border of renal tubules is highly enriched in dynamin 8 , a GTPase best known for its role in endocytosis 9 . Dynamin has an intrinsic propensity to assemble into multiple oligomerization states such as dimers, tetramers, rings, and spirals 9 . We identified for the first time direct dynamin-actin interactions 10 and showed that dynamin's oligomerization regulates actin polymerization in podocytes 11,12 , specialized cells essential for the selectivity of the kidney filter. Using murine models of CKD, we have shown that activation of dynamin-dependent actin polymerization reverses podocyte injury by restoring their unique structure and function 12 .
Here we show that in renal tubular epithelial cells dynamin cross-links filamentous actin (F-actin) into branched networks. Dynamin's cross-linking capability is defined by its oligomerization state and the length of F-actin. Pharmacological activation of dynamin oligomerization counteracts AKI by stabilizing the actin networks and thus cell integrity, which partially protects renal epithelial cells from oxidative stress-induced injury. Our study identifies the actomyosin cortex of the apical membrane of the renal tubular cell as a druggable target in AKI via dynamin as a proxy.

Results
Dynamin oligomerization establishes the stiffness and morphology of the apical membrane. To examine the role of dynamin-actin interactions in polarized renal tubular epithelial cells, we utilized Bis-T-23, an allosteric activator of actin-dependent dynamin oligomerization in a reconstituted system 13 , in the cells 11,13 , and in the whole organism 12 . Cellular phenotypes were assessed in Madin-Darby Canine Kidney (MDCK) cells by following the status of the F-actin and the staining pattern of a tight junction protein zonula occludens-1 (ZO-1), which is considered a biomarker of cell polarity. Cytochalasin D (CytoD) and latrunculin A (LatA), known inhibitors of actin polymerization, decreased F-actin levels, and induced discontinuous ZO-1 staining ( Supplementary Fig. 1a). In contrast, Bis-T-23 induced a slight increase in F-actin levels without any effect on ZO-1 staining. Addition of Bis-T-23 prior to but not after LatA, partially preserved F-actin levels and cell polarity. Neither DMSO vehicle nor dynamin inhibitor dynole 14 exhibited any effect ( Supplementary  Fig. 1a).
Scanning electron microscopy (SEM) allowed us to visualize drug-induced alterations of cell morphology focusing on the apical membrane (Fig. 1a). The average MDCK cell height was 11 ± 2 µm, and the average length of the microvilli was 0.63 ± 0.2 µm (Table 1), which is within the range observed in the kidney 15 . LatA decreased cell height, microvilli length and shifted the uniformly distributed microvilli into clusters, whereas Bis-T-23 induced the opposite effects (Table 1, Fig. 1a). When added prior to LatA, Bis-T-23 partially preserved cell height and microvilli length. Since microvilli exhibit exquisite length control defined by the cortical actin at their base 16 , these data provide evidence that Bis-T-23 modified the actomyosin cortex at the apical membrane.
To determine the exact effect that Bis-T-23 had on the cortical actin, we visualized the actomyosin cortex within the lamellipodia using platinum replica electron microscopy (PR-EM). LatA decreased the density of the actin networks, and this effect was partially abrogated by the addition of Bis-T-23 prior to LatA (Fig. 1b). As LatA accelerates actin filament depolymerization by sequestering actin monomers, we next examined whether the observed preservation of actomyosin cortex by Bis-T-23 was due to its positive effect on actin polymerization. In contrast to the potent stimulation of actin polymerization observed in podocyte cell extracts 10,11 , Bis-T-23 only marginally increased actin polymerization in MDCK cell extract ( Supplementary Fig. 1b). Similarly, immunodepletion of endogenous dynamin-2 (Dyn2) from the extract or inhibition of its GTPase activity by dynole resulted in marginal impairment of actin polymerization ( Supplementary Fig. 1c). While LatA and CytoD significantly impaired actin polymerization, the addition of Bis-T-23 prior to LatA or CytoD was not able to overcome their inhibitory effects ( Supplementary Fig. 1d, e). Jasplakinolide, a drug that induces actin polymerization by stimulating actin filament nucleation 17 , did not significantly increase the overall level of polymerization ( Supplementary Fig. 1d), suggesting that MDCK cell lysate exhibit a near maximal level of polymerized actin. Together, these data indicated that the effects of Bis-T-23 on the morphology of the apical membrane in MDCK cells were driven by a mechanism other than actin polymerization.
Given the common knowledge of enriched localization of Dyn2 and F-actin at the brush border of renal epithelial cells 8 , and dynamin's role in endocytosis, we next investigated if Bis-T-23 was affecting actin indirectly via alterations in endocytosis. As expected, both Dyn2 and F-actin co-localized at the actomyosin cortex underneath the apical membrane, within the microvilli, and at clathrin-coated pits (CCPs), defined by their distinct shape and size ( Supplementary Fig. 1f). We examined the dynamics of CCPs using total internal reflection fluorescence (TIRF) microscopy 18,19 . Bis-T-23, even at its highest concentration, had no effect on the distribution of the CCPs' lifetimes, whereas dynole decreased the number of productive CCPs (Supplementary Fig. 1g). This lack of correlation between the level of endocytosis and alterations in cell morphology asserts that Bis-T-23 targets the cortical actin without influencing dynamin's role in endocytosis.
Since renal cell polarity is maintained by the architecture and sustained contraction of the actomyosin networks, which establishes cell stiffness at the apical membrane 20 , we next measured cell stiffness using atomic force microscopy (AFM). Nanowizard IV system and JPK analysis software were used to determine changes in Young's Modulus 21 under different experimental settings ( Supplementary Fig. 2a). Treatment with LatA resulted in a significant decrease in cell-cell contact stiffness and apical cell stiffness in MDCK cells (Fig. 1c-e). In contrast, Bis-T-23 significantly increased cell stiffness compared to the DMSO vehicle (Fig. 1c-e), consistent with its positive effects on cell height, microvilli number, and the density of actin networks (Table 1 and Fig. 1b) 22 . The addition of Bis-T-23 prior to LatA strongly reduced the negative effect of LatA on cell stiffness ( Fig. 1c-e), in accordance with Bis-T-23's positive effect on actin networks and apical cell morphology (Table 1, Fig. 1b).
We have also determined cell stiffness using the BioScope II system as an alternative experimental approach for AFM. In this instance, force indentation curves were obtained following the model of Discher and coworkers computed with Matlab software 23 . Similar trends with regard to cell stiffness were recorded for the interplay between LatA and Bis-T-23 (Supplementary Fig. 2b, 2c). In addition, dynole did not exhibit any effect on cell stiffness, whereas CytoD significantly decreased cell stiffness ( Supplementary Fig. 2d, e), in accordance with their actin phenotypes ( Supplementary Fig. 1a). Together, these data establish the correlation between the status of the actomyosin cortex, cell stiffness, the morphology of the apical membrane, and cell polarity. These findings also convincingly demonstrate the role of dynamin oligomerization in defining mechanical parameters of epithelial cell polarity via its effect on the actomyosin cortex. b Representative PR-EM images of MDCK cells treated as explained in (a). The images show changes in the organization of the actomyosin cortex in MDCK cells under the indicated conditions. c Representative images of Young's Modulus maps of MDCK cells treated as explained in (a). d, e Bar graphs representing Young's Modulus depicting cell stiffness measured at the cell-cell junction (d) or at the apical membrane (e). Each symbol represents the average stiffness of a single cell. Results shown in d, e were generated from at least 10 cells from at least three culture dishes. Error bars, mean ± S.D. (*P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, ****P ≤ 0.0001, unpaired two-tailed t-test). ns, not significant. Dynamin cross-links actin filaments into branched networks that underlie cell polarity. In order to elucidate the molecular mechanism by which dynamin oligomerization influences the architecture of the actomyosin cortex, we next examined the effect of dynamin on actin filaments in a reconstituted system. Based on the current hypothesis, the length of actin filaments defines their mode of cross-linking 6 . As the average length of cortical actin filaments within a network at the leading edge is between 100 and 150 nm 24 , we examined the effects of dynamin on the organization of shorter filaments generated by capping F-actin with gelsolin (Gsn-actin) (Fig. 2a). The addition of Dyn2 resulted in the formation of large, branched networks (Fig. 2b, c). Based on the sizes and shapes of recombinant Dyn2 (Fig. 2d), the networks were formed predominantly by Dyn2 dimers (Dyn2 DIMER ) and tetramers (Dyn2 TETRA ) that interacted with several actin filaments ( Fig. 2e): Dyn2 DIMER bound up to two filaments, Dyn2 TETRA bound up to four filaments, and Dyn2 RING bound up to six filaments. Low magnification of the images revealed that dynamin-dependent networks form a pattern of smaller and larger ring-like shapes (Fig. 2c).
To correlate observations from the reconstituted system with dynamin's role in cells, we next determined the localization of endogenous Dyn2 on cortical actin networks using a monoclonal anti-Dyn2 antibody followed by a gold-conjugated secondary antibody ( Supplementary Fig. 3a-c). As seen in the reconstituted system, dynamin associated with a distinct number of F-actin within branched networks (Fig. 2f). Together, these data identify a novel activity of dynamin, that is cross-linking F-actin into branched networks.
To correlate dynamin's cross-linking capability and the protective effect of Bis-T-23 on the actomyosin cortex and the morphology of the apical membrane, we next examined the effects of Bis-T-23 on dynamin-mediated networks in reconstituted systems (Fig. 3a). Based on contour plots, which provide topographical representations of varying filament densities, Bis-T-23 increased overall network density (Fig. 3a)  , tetrameric (orange), partial/full ring (green). e High magnification images of actin filaments (pink) associated with distinct Dyn2 oligomerization forms (yellow). Insets show images of only Dyn2 at different oligomerization states. f Immunogold platinum replica electron micrographs focusing on actomyosin cortex in MDCK cells. Cellular localization of endogenous Dyn2 was determined by monoclonal Dyn2 antibody and gold-conjugated secondary antibody (white particles). To better visualize the gold particles within tightly packed actin networks, white densities associated with gold particles were pseudo-colored red, and the actin filaments associated with gold particles were pseudo-colored pink. The presence of multiple gold particles identifies the formation of macromolecular dynamin complexes on actin filaments, consistent with the formation of dynamin rings. Representative images of two independent experiments. explained by the increase in the number of F-actin bound to dynamin due to the increase in its oligomerization. In addition, dynamin more potently cross-linked shorter filaments than long F-actin ( Fig. 3a-c), suggesting that dynamin's capability to form branched networks is defined by its oligomerization status and the length of actin filaments. The ability to cross-link actin filaments into networks was shared by two dynamin isoforms, ubiquitously expressed Dyn2 and neuron-specific dynamin-1 (Dyn1) (Fig. 3b, c). We and others have shown that dynamin binds F-actin via two distinct domains: one situated in the Middle domain 10,12,[25][26][27] , and one part of the C-terminal proline-arginine rich domain (PRD) 28 . Similar networks were formed by Dyn1 and Dyn1 ΔPRD ( Supplementary Fig. 3d), demonstrating that PRD is dispensable for network formation. Finally, Dyn2 also cross-linked F-actin into tight bundles and hyper-bundles ( Supplementary Fig. 4a, 4b), as seen before 10,28 . In contrast to network formation, dynamin's ability to form bundles is dependent on PRD-actin interactions 28 . Bis-T-23 promoted the formation of hyper-bundles (Supplementary Fig. 4c, 4d), suggesting a mechanism by which Bis-T-23 increased the number of microvilli on the apical membrane (Table 1). This hypothesis is supported by the presence of antidynamin antibodies within actin bundles that underlie filopodia ( Supplementary Fig. 4e). Together, these data show that dynamin's ability to cross-link filaments into networks and bundles is influenced by its propensity to oligomerize as well as the length of actin filaments ( Supplementary Fig. 4f).
Insights generated by the reconstituted system were tested in the cell-based assays. Expression of both dynamin isoforms lacking PRD (Dyn ΔPRD ) in MDCK cells did not affect the ZO-1 staining pattern when compared to wild-type dynamins, and Dyn2 ΔPRD also increased F-actin levels ( Supplementary Fig. 5a). These data suggested that dynamin's ability to form tight bundles does not contribute to the overall level of F-actin or the cell polarity in MDCK cells. In contrast, expression of Dyn2 K/E , mutant impaired in F-actin binding 10 , decreased F-actin levels and cell polarity compared to Dyn2 expressing cells (Supplementary Fig. 5a-c), implicating the Middle domain in the formation of actomyosin networks that underlie cell polarity. This hypothesis is further supported by AFM experiments showing that Dyn2 K/E also decreased cell stiffness at the apical membrane when compared to Dyn1 WT ( Supplementary Fig. 5d).

Gsn-actin + Dyn2
Gsn-actin + Dyn2 + Bis-T-23 a F-actin + Dyn2 F-actin The inset shows the representative size (parameter) of actin networks for each condition. 11 to 23 actin networks were analyzed. c Graph depicting the density of the actin filaments incorporated into actin networks (blue) or loosely distributed through the background of the micrograph (yellow). Data were generated by measuring density within a defined square (100 × 100 nm) as shown in the representative insert. Results shown were generated based on EM data shown in (a). 33-45 dense networks (blue squares) and 14-27 loose networks (yellow squares) were counted per condition. Data in (b), (c) are plotted as mean ± S.E.M. and mean ± S.D, respectively (****P < 0.0001, one-way ANOVA with Tukey's multiple comparison test). ns, not significant. These studies are complementary to experiments using mouse inner medullary collecting duct (mIMCD) cells, another cell line representing polarized renal epithelial cells. In Dyn2 knockdown mIMCD cells, a significant decrease in F-actin levels and a patchy ZO-1-staining pattern were observed ( Supplementary Fig. 6a-c). The F-actin phenotype was rescued to normal level by the expression of either wild type or Dyn2 ΔPRD , but not by Bis-T-23 (Supplementary Fig. 6d-g). Together these data provide evidence for the essential role of dynamin and its oligomerization in the establishment and maintenance of epithelial cell polarity. They also strongly suggest that actin networks that underlie the apical membrane are formed by interactions between F-actin and the Middle domain, and that dynamin's ability to form bundles and hyper-bundles is not essential for cell polarity.

Increase in crosslinking capability
Pharmacological stabilization of cortical actin via dynamin ameliorates AKI. Loss of renal tubules brush border due to disassembly of actomyosin cortex is a predominant feature of AKI 2,29 . A widely used anti-cancer drug often leading to AKI is cisplatin 30 . A pivotal consequence of cisplatin injury in renal cells is an increase in reactive oxygen species (ROS) level 31 . Moreover, increased levels of ROS lead to a decrease in actin dynamics 32 , which causes disruption of the mitochondrial membrane; this further exacerbates ROS production 33,34 . Thus, we examined whether pharmacological stimulation of dynamin's cross-linking capability is sufficient to counteract cisplatininduced injury.
As seen before 35 , cisplatin decreased MDCK cell height, microvilli number, and length, and affected cell polarity without affecting endocytosis (Table 1, Supplementary Fig. 7a-d). TEM analysis suggested that these phenotypes could be attributed to cisplatin-induced loss of F-actin (Fig. 4a). The addition of Bis-T-23 prior to cisplatin abrogated its effects by stabilizing the actomyosin cortex against cisplatin-induced injury (Fig. 4a-c). Similar phenotypes were observed in porcine kidney proximal tubule (LLC-PK1) cells ( Supplementary Fig. 8). Together, these data provide evidence that Bis-T-23 preserves the actomyosin cortex from cisplatin-induced disassembly.
In a complementary physiologically relevant approach 36 , Bis-T-23 counteracted cisplatin's ability to reduce transepithelial electrical resistance (TER) as measured across the live epithelial monolayer using a transwell culture system (Fig. 4d). Since TER measures the integrity of tight junctions in cell culture, these data further establish the beneficial effect of Bis-T-23 on epithelial cell integrity upon injury.
As altered actin dynamics can further enhance ROS production, we next examined whether stabilization of the actin network by Bis-T-23 can counteract oxidative stress in cisplatin-injured cells. We elected to assess the status of cellular biomolecule carbonyls, a stable biomarker of oxidative damage produced by high levels of ROS, using our recently developed fluorescent sensor TFCH 37 (Fig. 4e). Indeed, the addition of Bis-T-23 prior to cisplatin partially diminished cisplatin-induced increase in carbonylation (Fig. 4f). Similar trends regarding carbonylation were observed after treatment with Bis-T-23 and the actin depolymerizer LatA (Supplementary Fig. 9a). Together these data provide evidence that stabilization of the actin networks can counteract the feedback loop between oxidative stress and the status of actin in injured cells.
An ex vivo rat kidney model was next utilized to visualize Bis-T-23-driven preservation of the actomyosin cortex at the brush border. Cisplatin induced a significant loss of F-actin staining at the brush border of proximal tubules (Fig. 4g). A similar loss of F-actin at the brush border upon the onset of severe ischemiareperfusion injury was recently observed using intraviral imaging 38 . Pre-treatment with Bis-T-23 partially protected cisplatin-induced loss of F-actin at the brush border of renal tubules (Fig. 4g), further demonstrating the effect of Bis-T-23 on actomyosin cortex at the apical membrane of renal epithelial cells. As seen in the cell culture, Bis-T-23 did not affect endocytosis in the tissue slices ( Supplementary Fig. 9b), further demonstrating that Bis-T-23 does not affect dynamin's role in endocytosis in renal proximal tubules.
Finally, we examined the reno-protective effect of Bis-T-23 in a mouse model of cisplatin-induced AKI 39 . BL6 mice were injected with either Bis-T-23 or DMSO once per day starting 24 h prior to injection of cisplatin. As seen before 40 , serum creatinine (SCr) and blood urine nitrogen (BUN) levels significantly increased 3 days post cisplatin injection (Fig. 4h). DMSO exhibited mild reno-protection as seen before 41 , due to its oxidant scavenging ability 42 . Daily administration of Bis-T-23 outperformed DMSO with regard to reno-protection (Fig. 4h). As all animals were sacrificed on day 5 due to the systemic toxicity of cisplatin ( Supplementary Fig. 9c), we could not examine the long-term benefits of Bis-T-23 on kidney function. Together, these studies persuasively demonstrate that preserving renal cell integrity via stabilization of the actomyosin cortex at the apical membrane counteracts cisplatin-induced nephrotoxicity.
Stabilization of the actin networks counteracts iohexol-induced AKI. While the exact mechanism of contrast dye-induced AKI is not fully deciphered, it remains a prime cause of hospitalacquired AKI 43 . Physiologically, contrast dye increases the osmotic load, decreases renal blood flow, and induces renal arterial constriction 44 . At the cellular level, ROS is a critical player in contrast to dye-induced AKI 45 . Given the role of ROS in regulating the actin cytoskeleton and our hypothesis that stabilization of the actomyosin cortex can counteract oxidative damage, we examined the effects of Bis-T-23 on a contrast dye-, iohexol-induced AKI 46 .
The cellular effects of iohexol were examined in human kidney proximal tubular (HK-2) cells. As seen for cisplatin, iohexol induced loss of F-actin, which was accompanied by a significant increase in the level of carbonylated molecules, an irreversible consequence of elevated ROS levels (Fig. 5a, b). Both phenotypes were partially counteracted by pre-treatment with Bis-T-23, further establishing the synergy between the status of the actin cytoskeleton and the level of oxidative stress in these cells. Complementary to cell-based assays, daily administration of Bis-T-23 preserved renal function based on the Scr and BUN levels in BL6 mice challenged with contrast dye (Fig. 5c). A much-coveted response of improved survival rate was also observed in the animals that received Bis-T-23 along with the contrast dye (Fig. 5d).
We have recently shown that elevated serum levels of soluble urokinase-type plasminogen activator receptor (suPAR) sensitize the kidney to iohexol-induced AKI 47 . We next investigated if dynamin-mediated protection of the tubular cells can also render reno-protection in BL6 mice expressing high levels of suPAR from fat tissue (suPAR-Tg) 48 . As expected 47 , the suPAR-Tg mice exhibited a decrease in kidney function 24-48 h post iohexol injection (Fig. 5e). In contrast, the animals that received a daily dose of Bis-T-23 exhibited significantly better kidney function despite the contrast dye challenge (Fig. 5e). Notably, a significant decrease in the mortality rate was observed in the mice cohort that received Bis-T-23 (Fig. 5f). As anticipated, the extent of oxidative damage (biomolecule carbonylation) in HK-2 cells concomitantly treated with Bis-T-23 was significantly lower than in cells that received only a combination of suPAR and Iohexol (Supplementary Fig. 9d).
In contrast to ROS-mediated cellular injury caused by cisplatin and iohexol, suPAR promotes tubular cell injury partly by increasing oxygen consumption rate (OCR) 47 . To link the physiological effect of Bis-T-23 in suPAR transgenic mice to its effect on cell metabolism, we examined whether stabilization of the actin network via dynamin decreases suPAR-driven OCR. OCR was measured in real-time in HK-2 cells under basal conditions and in response to sequential injections of mitochondrial inhibitors using a Seahorse XFe24 extracellular flux analyzer (Fig. 5g). The addition of an anti-suPAR antibody or Bis-T-23 ameliorated suPAR-driven increases in mitochondrial basal respiration, ATP production, maximum rate of respiration, and spare-respiratory capacity (Fig. 5f). These data demonstrate the positive effects of Bis-T-23 on cell metabolism upon injury. Together, these studies further affirm the feasibility of protecting against multiple types of AKI and thereby improving the survival rate in rodent models via dynamin as a proxy.
Simultaneous stabilization of actin in distinct kidney cells attenuates nephron injury. We have previously shown that pharmacological activation of dynamin-driven actin polymerization restored the structure and function of podocytes in diverse murine models of CKD 12 . In this study, we demonstrated dynamin's ability to preserve the integrity of tubular cells upon acute injury by cross-linking the actin cytoskeleton. Together, these insights led us to envision the possibility of simultaneously counteracting both, glomerular and tubular injury, by pharmacologically targeting dynamin.
To test this hypothesis, we examined whether Bis-T-23 could delay the loss of kidney function in the mouse model of Alport syndrome (AS). AS is an inherited form of progressive kidney failure due to mutations in the COL4A3, COL4A4, or COL4A5 genes that together encode type IV collagen, a major component of the glomerular basement membrane (GBM) 49,50 . Although the primary defect in AS is foot process effacement and proteinuria due to alteration in the composition of the GBM, the tubular response is also critical to the pathogenesis of AS and may precede the deterioration of glomerular function [51][52][53] . Indeed, signaling between tubules and glomerulus influences the response of the podocytes to collagen deficiency 54 . Currently, there is no specific therapy for AS.
Daily administration of Bis-T-23 slowed down the increase in proteinuria in Col4a3 −/− animals 55 (Fig. 6a), demonstrating that dynamin-mediated increase in actin polymerization in podocytes can counteract even genetic defects in GBM. Bis-T-23 also slowed down the decrease in kidney function, indicated by a small increase in BUN levels, protected tubules from developing hyaline ARTICLE NATURE COMMUNICATIONS | https://doi.org/10.1038/s41467-022-30101-4 casts, and extended the lifespan of animals (Fig. 6a-c). These data further attest to the positive impact of increased dynamin crosslinking activity on the preservation of renal tubular cell integrity. Together, our study demonstrates the suitability and the unique advantage of pharmacological targeting of nephron as a unit via dynamin as a proxy (Fig. 6d).

Discussion
Since the identification of direct dynamin-actin interactions 10 , a growing body of evidence is establishing this GTPase as one of the major regulators of the actin cytoskeleton in the cell. In contrast to canonical ABPs, the mechanisms by which dynamin influences actin are both, highly versatile and cell-type specific. This is due to the combination of dynamin's multiple oligomerization states and its ability to bind F-actin by two different binding sites. F-actin interactions with dynamin rings and helices via its C-terminal PRD result in actin bundles and hyper-bundles, which have been implicated in the formation of filopodia 56 and membrane protrusions during myoblast fusion 28 . Interactions between dynamin's Middle domain and F-actin have been implicated in regulating actin networks in lamellipodia 26 , postsynaptic cytoskeleton organization, and neuromuscular junction development 27 , actin bundle rigidity in invadosomes during myoblast fusion 25 , and formation of podocyte foot processes 12 . While the formation of foot processes is driven by dynamin-stimulated actin polymerization, this study suggests that the molecular mechanism by which dynamin influences these other actin-driven processes might be in part due to its ability to cross-link F-actin into branched networks. Our current study expands the role of dynamin-dependent network formation to include formation of actomyosin cortex at the apical membrane of polarized epithelial cells and the establishment of cell stiffness.
Despite the mechanistic diversity, dynamin's capability for cross-linking F-actin or promoting actin polymerization is enhanced by the increase in its oligomerization state (this study and refs. 12,28 ). Dynamin oligomerization is cooperative and is regulated by its concentration 57 , the length of actin filaments (this study and ref. 13 ), and SH3-domain-containing proteins 25,58 . The combination of all these mechanisms ultimately defines the temporal, spatial, and cell-type specificity of dynamin's role in modifying and/or establishing diverse actin structures.
Changes in mitochondrial function and cell metabolism have been linked to a multitude of AKI etiologies 59 . Mitochondria are dynamic organelles that respond to physiological signals and are significant sources of ROS in healthy 60 and injured cells 61 . Cisplatin and iohexol directly damage mitochondria leading to increased production of ROS 37,43,[62][63][64] , while suPAR affects the bioenergetic parameters of renal cells 47 . As many cytoskeletal proteins are sensitive to ROS 32,65 , a decrease in actin dynamics in the presence of elevated levels of ROS has been reported 32 . Meanwhile, the dysregulated actin cytoskeleton further augments ROS production 33 , suggesting a feedback loop between oxidative stress and the actin cytoskeleton. Here we show, by using a combination of cell-based assays focusing on cell polarity, realtime extracellular flux experiments, and our new carbonylationspecific fluorophore, that stabilization of actomyosin cortex via dynamin partially attenuates the feedback loop between oxidative stress and actin cytoskeleton dynamics regardless of the initial type of injury (cisplatin, LatA, iohexol, or suPAR).
It is well established that CKD and AKI, though mechanistically distinct, are closely interconnected, with AKI being recognized as a risk factor for CKD development and progression 66 . As of now, AKI remains undruggable 67 . This is particularly relevant in light of the current coronavirus disease 2019 (COVID-19) pandemic, where hospitalized patients with COVID-19 and an elevated suPAR plasma level develop AKI at alarming rates 68 . Elevated suPAR levels have been linked to both AKI 47 and CKD 69 , further reinforcing the molecular link between these two distinct kidney diseases. Therefore, it is becoming increasingly essential to develop therapeutics that can protect against an array of renal insults on multiple cell types 70 . Here we report the reno-protective effect of a dynamin agonist in a genetic model of AS, which exhibits injury to both podocytes and tubular cells. Our study establishes a comprehensive approach for developing novel therapeutics that engage with the actin network in multiple cell types within the nephron and treat myriad kidney diseases regardless of the site of injury.  Dynamin agonist counteracts Iohexol-induced AKI. a Status of F-actin and biomolecule carbonylation in HK-2 cells. HK-2 cells were treated with medium (Control), DMSO (0.1%), or Bis-T-23 (10 µM, 0.1% DMSO) for 1 h. The medium was replaced with only medium or medium containing Iohexol +DMSO (250 mg/mL (F-actin set) and 100 mg/mL (TFCH assay)) or iohexol+Bis-T-23 for~3 h. Scale bar, 20 μm. b Graphs depicting the relative levels of F-actin within a fixed region of interest (ROI) in the cells (58-98 ROI analyzed) TFCH fluorescence (cellular biomolecule carbonyls) (data points (n) = 46-83; each point represents average intensity of 1-4 cells). Error bars, mean ± S.D. (*P < 0.05, ***P < 0.001 ****P < 0.0001, one-way ANOVA with Tukey's multiple comparison test). c, e Scatter dot plots showing AKI induced by Iohexol (5 g/kg) determined by the level of SCr or BUN in BL6 wild-type mice (c, d) or suPAR-Tg mice (e, f). c, e Error bars, mean ± S.E.M. (P values are reported in the Figure, unpaired two-tailed t-test). As indicated, animals were injected with either DMSO (1%) or Bis-T-23 (20 mg/kg) once a day starting at 0 h. The measurements were performed at the indicated times. Number of animals per condition in c: BL6 control (n = 20), DMSO (n = 18), and Bis-T-23 (n = 21). Number of animals per condition in (e): suPAR-Tg control (n = 12 for BUN or SCr), DMSO (n = 19 for BUN; n = 16 for SCr), and Bis-T-23 (n = 20 for BUN; n = 13 for Scr). d, f Log-Rank (Mantel-cox) test was performed to analyze survival curves of animals treated as described in (c), (e). P was calculated using an unpaired two-tailed t-test. g Oxygen consumption rate (OCR) curves and Seahorse XF analyzer measurements of mitochondrial respiration of HK-2 cells. HK-2 cells were treated with suPAR (10 ng/ml), human anti-uPAR antibody (50 ng/ml), or Bis-T-23 (10 µM) alone or in combination for 24 h. OCR was measured in real-time under basal conditions and in response to sequential injections of mitochondrial inhibitors including oligomycin (OLG; an ATP synthase inhibitor), FCCP (an uncoupler of ATP synthesis from oxygen consumption), rotenone (ROT; complex I inhibitor), and antimycin A (AA; complex III inhibitor). Each OCR value was normalized to cell number and is presented as pm/min/100,000 cells.  10 or scrambled shRNA (Sigma-Aldrich) in complete media containing polybrene (8 µg/ml). After 24 h, the media was replaced every other day with media containing puromycin to select for stable transfectants. The extent of Dyn2 knock-down was assessed by western blot. All cell lines were grown on collagen-coated glass coverslips.
Proteins. Human Dyn1, rat Dyn2, human Dyn1ΔPRD, and rat Dyn2ΔPRD were expressed using Bac-to-Bac baculovirus expression systems (ThermoFisher Scientific) in Sf21 insect cells 10 . Recombinant human gelsolin (Gsn) (gift from Fumihiko Nakamura, Brigham and Women's Hospital); purified non-labeled G-actin, pyrene labeled G-actin (Cytoskeleton); human uPAR protein (R&D Systems); Rhodamine phalloidin (ThermoFisher Scientific).  Figure, unpaired two-tailed t-test). b Log-Rank (Mantel-cox) test was performed to analyze the survival curves of the two treatment groups. c Representative histopathology of kidney determined by periodic acid-Schiff (PAS) and H&E staining on samples obtained from Col4a3 −/− mice described in (a). Animals were 62 days old (day 20 of Bis-T-23 or DMSO treatment). Glomerulopathy in Col4a3 −/− DMSO-treated animals was characterized by mesangial expansion observed on the PAS staining (black arrow), increased cellularity, and an increase in Bowman's space. Degenerative changes in tubules included the formation of casts (*) and the presence of tubular sclerosis. Bis-T-23 treatment over 20 days partially preserved the morphology of glomeruli (diminished mesangial expansion) and tubules (absence of casts). d Schematics of the nephron (image credited to National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health) and the mechanism involved in dynamin regulated reno-protection. In polarized epithelial cells of renal tubules, dynamin establishes cell polarity by cross-linking actin filaments into networks (this study). Dynamin's ability to cross-link filaments into bundles is implicated in the formation of microvilli. Acute injury increases ROS production, alters cell metabolism, and perturbs the actin dynamics, which further enhances ROS production leading to more pronounced cellular oxidative damage (biomolecule carbonylation). An increase in dynamin's cross-linking capability in the actomyosin cortex preserves tubular cell integrity, which partially protects them from oxidative insult. Additionally, dynamin is essential for the structure and function of podocytes. Pharmacological activation of dynamin restores foot processes by inducing actin polymerization. Our study shows that pharmacological activation of dynamin oligomerization exhibits a dual beneficial effect on the nephron by targeting two distinct molecular mechanisms: preservation of the integrity of podocytes (actin polymerization) and renal tubular cells (cross-linking of actin filaments).
MDCK, mIMCD, mIMCD Dyn2 KD , HK-2 and LLC-PKI cell imaging, and quantification. All images were captured using a LSM 5 PASCAL or a 600 LSM (Zeiss). For image quantification, all images within an experimental set were acquired using identical optical parameters, thresholded, and analyzed using Fiji (Image J) or Image J (NIH) software. To measure the continuity of tight junctions, a free line was drawn along ZO-1 staining. The graph represents a sum of the length of ZO-1 staining as a percentage of the total length of the cell membrane. A defined region of interest (ROI) drawn with the rectangle tool on the ZO-1associated cell membrane was quantified to assess ZO-1 intensity. For MDCK cells, the intensity of the F-actin within each cell was quantified. For mIMCD or mIMCD-Dyn2 KD cells, the intensity of F-actin in a fixed ROI within each cell was quantified. Cellular carbonylation level was determined by quantifying TFCH fluorescence per cell 37 . In the scatter dot plots for TFCH assays, each dot represents the average intensity of 1-14 cells. The representative images showing cellular carnbonylation levels were processed identically within a set and assigned a pseudocolor for visual clarity. The staining intensity of treated samples was normalized to the non-treated or vehicle control. An unpaired two-tailed t-test or oneway ANOVA with Tukey's multiple comparisons test was performed using Prism (GraphPad). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001; whereas P > 0.05 was considered not significant.
Immunodepletion of endogenous Dyn2 from MDCK cells. Cytosolic extracts were incubated with anti-Dyn2 antibodies (1 μg/200 μg cytosolic proteins) at 4˚C for 2 h, followed by incubation with protein G PLUS-agarose beads at 4˚C for 2 h. After centrifugation, the supernatant was used as the ΔDyn cytosol for the actin polymerization assay. Mock-depletion was performed in the same manner with a non-specific mouse anti-rabbit IgG antibody (1 μg/200 μg cytosolic proteins). The efficacy of the depletion was confirmed to be >90% by Western blot using the same antibody.
Cell polarity assessment using a transwell system. MDCK cells were grown in collagen-coated transwells (Corning) for 48 h before treatment with DMSO (0.1%) or Bis-T-23 (30 µM; 0.1% DMSO) for 1 h and cisplatin (50 µM) or media for another 23 h. A Millicell ERS Voltohmmeter was used to measure the transepithelial electrical resistance (before and after 24 h of initiating the treatment) in each sample.
NanoWizard IV system. AFM imaging was performed using a NanoWizard IV (JPK Instruments) mounted on an Axiovert 200 inverted light microscope (Carl Zeiss). Microscopes were equipped with a heated stage (37°C) and cells were imaged in HEPES-buffered medium. As previously reported for MDCK cells, indentation measurements were performed with silicon nitride cantilevers (MLCT, Bruker) with a nominal force constant of 0.01 N/m 21 . Automated calibration was performed in contact-free mode as provided by the calibration manager in the JPK SMP software before every series of experiments. Areas of 2500 µm 2 were scanned in QI imaging mode to acquire 3600 independent force measurements per area. Cells were indented up to a force of 0.9 nN at a vertical speed of 18 µm/s. Determination of Young's Modulus was performed using JPK data analysis software. Fit parameters were set to the Hertzian model according to Sneddon. Cantilever was specified as quadratic pyramid tip shape with a half-angle to edge of 15°. Image analysis and quantification of apical stiffness and stiffness at cell-cell contacts were performed using Fiji software. Topography maps of confluent MDCK cells show elevation at cell-cell junctions. Cell contact sites were marked as ROI manually in the grayscale topography map and ROIs were transferred to the associated force map to analyze stiffness at the respective sites. For determination of apical cell stiffness, grayscale topography maps were analyzed using Fiji threshold presets MaxEntropy/Dark background to mark cells and identify ROIs. ROIs were transferred to the associated force map to determine cell stiffness. Data from at least 10 cells in at least three culture dishes were included in the analysis. Statistical analysis was performed as univariate analysis, defining a p-value of <0.05 as statistically significant. Bioscope II AFM system. AFM imaging was performed using a Bioscope II AFM (Veeco) mounted on an Olympus IX 73 inverted light microscope (Olympus). The microscope was equipped with a heated stage (37°C) and cells were imaged in HEPES-buffered medium. Measurements were performed in contact mode using silicon nitride cantilevers with a nominal force constant of 0.01 N/m. Cantilevers were calibrated before every series of experiments by thermal tune using the manufacturer's software. Force indentation curves were analyzed according to the model of Discher et al. 23 computed with MATLab software. Data from at least 10 cells in at least three culture dishes were included in the analysis. Statistical analysis was performed as univariate analysis, defining a p-value of <0.05 as statistically significant.
Electron microscopy (EM). All EM images were acquired at the Massachusetts General Hospital (MGH), Harvard Medical School, or Brandeis University EM core facility.
Scanning electron microscopy (SEM). MDCK cells were treated as described in the "Immunocytochemistry for ZO-1 and F-actin" section, fixed for 2 h in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4 (Electron Microscopy Sciences, EMS), rinsed with 0.1 M sodium cacodylate buffer, treated with 1% osmium tetroxide, rinsed in 0.1 M sodium cacodylate buffer and dehydrated through a graded series of ethanol reaching 100%. The samples were dehydrated in the critical point dryer (CPD) Autosamdri®-815 (Tousimis), coated with a 5 nm layer of platinum using a sputter coater (SC) EM ACE600 (Leica), and mounted on stubs using double-sided carbon conductive tape (EMS). Samples were examined at 3-5 kV using S-4700 FE-SEM (Hitachi). Cell height and microvilli length were measured using SEM software and ImageJ, respectively. Microvilli density in a constant area (0.7 in 2 ) on the membrane (≥10 areas per condition) was counted manually.
Transmission electron microscopy (TEM) experiments. (1) Negative staining of proteins in the reconstituted system: G-actin, F-actin and gelsolin-actin filaments (gelsolin:actin = 1:50) were prepared as described previously 10 . Gelsolin-actin was diluted to 10 μM and incubated with 2 µM recombinant Dyn1, Dyn2, or Dyn1ΔPRD for 60 min at 25°C in the presence or absence of Bis-T-23 (0.4 μM). All samples were diluted to 0.2 µM final actin concentration, and absorbed to glow discharged formvar-carbon coated copper grids for 1 min, blotted to remove excess solution, negatively stained with 1.5% (w/v) uranyl acetate for 1 min, blotted again, and allowed to air-dry. Images were captured at an acceleration voltage of 80 kV using an FEI Morgani 268 or JEOL JEM 1011 transmission electron microscope (TEM) equipped with CCD camera. The brush tool with 50% opacity in Photoshop (Adobe), and "contour plotter" in ImageJ were used for data presentation.
(2) Platinum replica EM and immunogold EM in cells: MDCK cells were treated as described in the "Immunocytochemistry for ZO-1 and F-actin" section, detergent extracted and processed for PR-EM as described previously 71 . For dynamin localization, samples were fixed with 0.2% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, quenched with NaBH 4 for 10 min, treated with Dyn2 antibody (1:100), and subsequently with 18 nm colloidal gold-secondary antibody (1:5). Cells were fixed and processed the same for PREM 71 . For PR-EM and immunogold PREM, samples were dehydrated in a CPD, coated with a 2 nm layer of platinum, and stabilized with 5 nm of carbon using SC. Samples were examined as described above and images were presented as inverted contrast.
Measurement of cellular bioenergetics. A Seahorse Extracellular Flux (XFe24) Analyzer (Agilent) was used to measure oxygen consumption rates (OCR) in real time 47 . HK-2 cells were treated with only media (control) or media containing human recombinant uPAR (10 ng/ml), human anti-uPAR antibody (50 ng/ml), or Bis-T-23 (10 µM) either alone or in combination for 24 h. Respiratory parameters were quantified, using the Cell Mito Stress Test Kit (Agilent), by subtracting respiration rates at times before and after the addition of mitochondrial activators and inhibitors 47 . Experiments were replicated in five wells, averaged for each treatment group and the data were presented as means ± standard errors of the means (SEM). One-way ANOVA using Prism was used to compare controls (CTL) with treatment groups (*P < 0.05, **P < 0.01, ***P < 0.001).
Rat kidney tissue slice experiment. Male Spraque-Dawley rats (strain code 400) were from Charles River. Rat kidney were prepared with minor modifications 29 . The slices were treated with DMSO (0.1%) or Bis-T-23 (30 μM, 0.1% DMSO) for 1 h before adding cisplatin (200 μM) for 8 h and fixing with periodate-lysine paraformaldehyde overnight at 4°C. For the albumin uptake assay, kidney slices were incubated in the presence or absence of Bis-T-23 (30 μM) for 1.5 h before adding rhodamine-albumin (100 μg/ml) for 30 min. Slices were then rinsed with PBS, fixed, and processed. Animal experiments were approved by the institutional committee on Research Animal Care at MGH, in accordance with the NIH Guide for the care and use of the laboratory animals (protocol #2012N000004 (PI, Brown)).